The bacterium Bacillus thuringiensis (Bt) contains genes encoding insecticidal proteins. Bt proteins are toxic when ingested by susceptible insect larvae. The protein attacks the insect's midgut, causes cessation of feeding, and eventually kills the insect. Bt toxins have been produced as fermentation products of Bt cultures and used in spray formulations for crop protection. Bt genes have also been used commercially to transform crop plants; these transgenic crop plants' cells then produce the insecticidal protein which attacks susceptible insects that attempt to feed on the plant.
The general mode of action of Bt toxins is well known in the art and is described for example by Rajamohan F, Lee M K, Dean D H (1998) Progress in Nucleic Acid Research and Molecular Biology 60: 1–27. The protein produced by the bacterium is usually a protoxin, which itself is not toxic until it is proteolytically cleaved by the insect's own proteases. The smaller protein resulting from proteolysis is the active toxin. This toxin diffuses through the peritrophic membrane to the midgut epithelium, where it binds to one or more sites in the membrane. This initial binding step may be reversible, but eventually the toxin becomes irreversibly bound to the membrane. A conformational change occurs in the toxin, whereby membrane-spanning alpha helices are inserted into the membrane, where they aggregate and form pores. These pores disrupt the normal osmotic balance of the epithelial cells. The cells swell and lyse, leading to destruction of the midgut epithelial cell layer and eventual death of the insect.
The initial binding step is believed to be necessary for toxin action; consequently there have been many studies on binding interactions of Bt toxins and components of the midgut, described for example by Pietrantonio P V and Gill S S (1996) in Biology of the Insect Midgut, Chapman & Hall, London, pp 345–372. Techniques used to study binding often start with the isolation of a brush border membrane vesicles (BBMVs) from the microvillar portion of columnar epithelial cells. Binding to BBMVs in suspension can be measured using labeled toxin. Alternatively, proteins can be isolated from BBMVs, separated by denaturing electrophoresis conditions, transferred to membranes, and probed with toxin. In addition, histological sections of insect midguts can be prepared and binding of labeled toxin can be visualized using microscopy.
Binding of Bt toxins to specific insect proteins can also be measured. Several proteins that interact with Bt toxins are well known in the art. Aminopeptidases exist in many different forms in insect midguts, and many of them have been shown to bind Bt toxins (Knight P J K, Knowles B H, Ellar D J (1995) Journal of Biological Chemistry 270 (30): 17765–17770; Gill S S, Cowles E A, Francis V (1995) Journal of Biological Chemistry 270 (45): 27277–27282; Luo K, Sangadala S, Masson L, Mazza A, Brousseau R, Adang M J (1997) Insect Biochemistry and Molecular Biology 27 (8-9): 735–743). Members of the cadherin superfamily have also been shown to bind Bt toxins (Vadlamudi R K, Weber E, Ji I H, Ji T H, and Bulla L A (1995) Journal of Biological Chemistry 270: 5490–5494; and Nagamatsu Y, Koike T, Sasaki K, Yoshimoto A, Furukawa Y, (1999) FEBS Letters 460: 385–390). Phosphatase enzymes have also been implicated in Bt toxin binding (Sangadala S, Walters F S, English L H, Adang M J, (1994) Journal of Biological Chemistry 269 (13): 10088–10092). TPP-75, an elastase-like serine protease, binds to certain Bt toxins and causes them to precipitate (Milne R E, Pang A S D, Kaplan H (1995) Insect Biochemistry and Molecular Biology 25 (10): 1101–1114). BTR-270, a peptidoglycan, binds Cry1A toxins with high affinity (Valaitis A P, Jenkins J L, Lee M K, Dean D H, Garner K J (2001) Archives of Insect Biochemistry and Physiology 46 (4): 186–200). Bt toxins have also been shown to bind to nonprotein components of midgut epithelial membranes. Glycolipids from Manduca sexta have been shown to bind Cry1A toxins using an overlay technique (Garczynski S F and Adang M J (2000) in Entomopathogenic Bacteria: From Laboratory to Field Application, Kluwer Academic Publishers, pp 181–197). Neutral lipids are involved in Bt toxin binding to Manduca sexta brush border membranes (Sangadala S, Azadi P, Carlson R, Adang M J (2001) Insect Biochemistry and Molecular Biology 32 (1): 97–107). Neutral glycolipids, especially hexa- and tri-saccharylceramides, are implicated in Cry1A toxin binding in diamondback moth (Kumaraswami N S, Maruyama T, Kurabe S, Kishimoto T, Mitsui T, Hori H, (2001) Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology 129 (1): 173–183).
The relationship between binding targets for Bt-toxins and susceptibility or resistance to Bt is very complicated and not completely understood at the present time. Several hundred strains of Bacillus thuringiensis exist, with considerable specificity toward various groups of insects. Co-evolution between the insects and Bt has resulted in specificity of the interaction between Bt-toxin and the membranes of insect gut cells. The Bt-toxin of a particular strain of Bacillus thuringiensis may bind to the gut of some insect larvae but not to others. Thus, the Bt-toxins may have a high specificity for a small number of insect pest species while having no significant activity against beneficial insects, wildlife, or humans.
Plants transformed to carry Bt genes and express insecticidal proteins are known in the art and include potato, cotton, tomato, corn, tobacco, lettuce, and canola. Transformed plants are known in the art as reflected in U.S. Pat. Nos. 5,608,142; 5,495,071; 5,349,124; and 5,254,799, the specifications of which are incorporated in their entirety herein by reference. The use of genetically engineered plants is designed to reduce the use of broad spectrum insecticides.
There is concern that resistance may evolve to Bt toxins, whether they are applied to plants in spray formulations or the plants are genetically engineered to express them. The development of resistance to Bt-toxin expressing crops may also result in resistance to commercial formulations of fermented strains of Bt such as DIPEL® (Abbott Laboratories).
Rapid, reliable methods for broad screening to distinguish and detect the development of Bt resistance in populations of insects are needed. Heretofore, all methods require living or fresh-frozen insect larvae or preparations derived from them. The simplest methods employ bioassays on living insects, in which survivorship or larval metabolic rates are determined over time following a diet containing a specified concentration of a Bt-toxin. One such bioassay based on reduced metabolic rates after exposure to low doses of toxin mixed into artificial diet is discussed in U.S. Pat. No. 6,060,039 to Roe et al. which is incorporated herein by reference. Other bioassays are based on survival after exposure to a single, high diagnostic dose of toxin (for example, Diaz-Gomez O, Rodriguez J C, Shelton A M, Lagunes-T A, Bujanos-M R, (2000) Journal of Economic Entomology 93 (3): 963–970).
In principle, these bioassay methods can detect resistance no matter what its biochemical or physiological mechanism is. However, they require living, healthy larvae for their use, which are not always available. A more severe limitation on these methods is that, depending on the frequency of resistance genes in the populations, millions of individuals may need to be tested to detect a single resistant larva. High-level resistance to Bt is usually recessive, which means that an insect must have two copies of the resistance gene to be resistant. To a very good approximation, the frequency of such homozygous individuals is given by the square of the frequency of the resistance allele. For example, if the resistance allele frequency is one in a thousand, the frequency of homozygous resistant individuals is one in a million. In this example, more than a million larvae would need to be screened to detect resistance.
One solution to this problem is to develop methods for detecting the resistance genes directly. In the example just given, the frequency of heterozygous carriers of one copy of the resistance allele is 2×0.001×0.999 or approximately 2 in a thousand. When resistance is recessive, these individuals would not be identified by bioassay because the one resistance allele they carry is not enough to make them fully resistant. But a direct, DNA-based method for detecting the resistance allele would identify these individuals, and sample sizes on the order of a thousand, rather than a million, would suffice.
The main limitation to developing DNA-based methods for detecting resistance alleles is that, up to now, the identity of resistance-causing genes has been unknown. In spite of much work on Bt toxin mode of action, prior to the invention described herein there has not been a demonstration of which genes, when mutated, actually cause resistance. Accordingly, there is room for variation and improvement in the art of screening assays useful in detecting the presence of genes conferring Bt resistance in natural populations.